Vol. 126, No. 1 Printed in U.S.A.
JOURNAL OF BACTERIOLOGY, Apr. 1976, p. 417-428 Copyright ©D 1976 American Society for Microbiology
Study on Development of Agaricus bisporus by Fluorescent Microscopy and Scanning Electron Microscopy K. N. SAKSENA,* R. MARINO, M. N. HALLER, AND P. A. LEMKE Carnegie-Mellon Institute of Research, Carnegie-Mellon University, Pittsburgh, Pennsylvania 15213
Received for publication 30 December 1975
Two strains of Agaricus bisporus have been investigated by fluorescent microscopy and scanning electron microscopy. Somatic nuclei, stained with auramin 0 and examined by fluorescent microscopy, appear to be randomly distributed, divide asynchronously, and assume a nonclassical or "two-track" configuration during mitotic metaphase. A similar configuration has been observed for nuclei during meiosis, but early meiosis in A. bisporus appears to be classical, usually with nine pairs of chromosomes evident during prophase I. Scanning electron microscopy has been used to document developmental stages in the formation and germination of basidiospores. Two-spored basidia were predominant, but occasionally one- or three-spored forms were observed. Fourspored basidia were absent, and uninucleated basidiospores were exceedingly rare to absent.
Studies on the cytology and genetics ofAgaricus bisporus (Lange) Imbach have shown consistently that the sexual cycle of this fungus is highly disorganized and atypical of a basidiomycete. The fungus has no semblance of dikaryotic organization, and the basidiospores, produced mostly two per basidium, contain an indefinite number of nuclei and represent predominantly self-fertile progeny (3, 5, 6, 13, 1517, 19, 20, 26, 27, 31, 32, 35, 40-45). We have examined the cytology and development of two strains of A. bisporus. Critical examination of these features has been possible through recent advances in fluorescent microscopy of fungal nuclei (18, 20) and through improvements in the preparation of biological specimens for scanning electron microscopy (2, 9, 10, 23, 24, 28, 30). MATERIALS AND METHODS Strains and media. Strains of A. bisporus designated D-26 and 70-2 were investigated. These strains represent, respectively, golden-white and light-brown varieties of the cultivated mushroom. Cultures were grown on a defined medium containing: asparagine, 1 g; glucose, 20 g; MgSO4 7 H20, 0.5 g; K2HPO4, 1 g; KH2PO4, 0.46 g; CaSO4, 0.5 g; FeCl3, 10 mg; thiamine, 120 Ag; distilled water, 1,000 ml; and agar, 20 g. Mushrooms were obtained commercially (Butler County Mushroom Farm, Inc., Worthington, Pa.) or were grown in the laboratory, using cased rye grain as a substrate (39). Basidiospores were collected as spore prints at 20 C and germinated either in the presence of isoamyl alcohol or isovaleraldehyde (1 and 0.1%, respectively, added to defined medium after autoclaving) or in the prox-
imity of a growing mycelium (23, 37, 47). Spores and mycelia were generally inoculated onto 5 cm2 membranes (DuPont PD 150 film) spread upon the surface of the defined medium. Cultures were incubated at 25 C, and membranes with adherent cultures were removed from the medium and prepared for microscope examination. Fluorescent microscopy. Procedures for examination of fungal nuclei by fluorescent Feulgen staining were described previously (20). Two dyes, auramin 0 (no. 10-120, Chroma-Gesellshaft, Stuttgart) and acrinol (no. Dl 660-6, Aldrich, Milwaukee), were used. Acrinol was used only to stain nuclei of basidial cells. Gill tissue was treated with Newcomer fixative (29) for at least 6 h and stored in 70% isopropanol prior to staining with the acrinol. In all other instances, fixation of cells was avoided and auramin 0 was used as the stain. Cells were placed in 1 N HCl at 60 C for at least 2 min prior to staining. Staining solutions were freshly prepared and contained 20 ml of water, 0.1 g of auramin 0 (or 0.1 g of acrinol), 0.2 g of potassium metabisulfite (0.2 g of sodium metabisulfite substituted if acrinol used), and 2 ml of 1 N HCl. Cells were stained for 20 min and then washed repeatedly in water and finally in 45% acetic acid. In each step of the procedure, cells were recovered from suspension by centrifugation (1,000 x g). Cells were placed on a microscope slide and air dried. The slide was then dipped momentarily in 95% ethanol, and material remaining on the slide was mounted for microscope examination in 65% sucrose (immersion oil substituted if acrinol used). A Leitz Orthoplan microscope fitted for incidentlight fluorescence with a Ploem vertical illuminator was used. The light source was an Osram superpressure mercury lamp (HBO 100 W/1 DC). During examination of material stained with auramin 0, the 117
418
SAKSENA ET AL.
microscope was set at position 3 with the following: one red suppression filter (BG38), two excitation filters (KP490), and one dichroic mirror (K490); no barrier filter was used. A X100 fluorite objective (1.39 numerical aperture) fitted with an iris (C. Reichert) was used. During examination of material stained with acrinol, the microscope was set at position 1 with the following: one excitation filter (UG1), one dichroic mirror (K400), and a barrier filter (K420). Either a X95 fluorite objective (1.32 numerical aperture) or a X90 apochromatic objective (1.40 numerical aperture) was used. Photographs were taken on Kodak Panatomic X film. Scanning electron microscopy. Segments of gill tissue (3 mm2) cut from fresh mushrooms at different stages of development were fixed with 6.25% glutaraldehyde in 0.05 M phosphate buffer (pH 7.0) for 3 h, washed repeatedly in buffer, and postfixed in 1% osmium tetraoxide for 1 h. After the washing, dehydration in a graded series of ethanol (30, 50, 70, and 90% for 15 min each, and 100% with three changes each of 15-min duration), and subsequent substitution in isoamylacetate for 15 min, critical point drying with liquid CO2 was performed with a Sorvall critical point dryer. Spore prints and cultures on membranes were air dried without fixation. Dried samples were coated with gold-palladium and examined with a JOELCO-JSM2 scanning electron microscope operating at 25 kV. The microscope was equipped with a nondispersive X-ray analyzer for elemental chemical analysis. Photographs were taken on Polaroid P/N55 film.
RESULTS AND DISCUSSION Initial experiments on the cytology of A. bisporus were conducted with basidiospores. These spores were collected as spore prints from opened mushrooms of two strains, D-26 and 702. Nuclei in these cells were stained with auramin 0 (Fig. 1A). The spores from the two strains proved to vary somewhat in nuclear distribution (Table 1). Uninucleated spores were observed rarely and only in one strain, 702. The majority of spores from this strain contained four nuclei, whereas the majority of spores derived from strain D-26 contained three or four nuclei. The surface of basidiospores examined by scanning electron microscopy proved to be smooth even at high (x 10,000) magnification (Fig. 2A). These dormant spores (47) can be stimulated to germinate at up to 50% frequency if placed in proximity to a growing mycelium of A. bisporus (22). Alternatively, spores can be induced to germinate by the addition of 1% isoamyl alcohol or 0.1 % isovaleraldehyde to the growth medium (37). Regardless of the method used to induce germination, spores do not germinate synchronously, and they produce a single germ tube per spore (Fig. 2B through F).
J. BACTERIOL.
The entire contents of a spore empty into this germ tube (Fig. 1B through E). Nuclei in mycelia do not divide synchronously or establish a dikaryon. Somatic nuclei are irregularly distributed and can be transferred to adjacent cells by hyphal anastomosis (Fig. 3A). Mature hyphae are often heavily encrusted (Fig. 3B and C). This encrustation, when observed by scanning electron microscopy, was resolved as crystals of at least three forms. X-ray analysis of crystals indicated the presence of potassium and calcium ions therein, and organic acids such as oxalate are assumed to be the major anionic component (21). Somatic nuclei of A. bisporus are often elongated (Fig. 4A) and assume a nonclassical configuration during mitotic division. This configuration has been observed in other higher fungi (1, 11, 12, 14, 38, 46) and has been called a "twotrack" configuration (4). Asynchrony of nuclei in hyphae of A. bisporus is evident in Fig. 4B, which shows a single, branched cell containing three types of nuclei: condensed, diffuse, and elongated ("two-track"). In A. bisporus the most common number of nuclei per vegetative cell is 3 (Fig. 4C through E), but as many as 25 nuclei per cell have been counted. Greater than 25 nuclei per cell have been seen in inflated (nonvegetative) cells found in the cap and stipe of mushrooms. Young mushrooms with normal morphology (Fig. 5A) have deep gills and a prominent veil. At maturity these mushrooms produce from one to several billion basidiospores. Abnormal mushrooms with reduced gill formation and scant veil tissue are sometimes found (Fig. 5B). In an extreme condition, asporogenic or imperfect mushrooms are formed (Fig. 5C), a condition that has been termed "open veil" (36). Basidia of A. bisporus develop asynchronously, and details of their meiosis are shown in Fig. 6 and 7. Nuclei and chromosomes in these cells were stained with acrinol. Early meiosis or prophase I appears to be classical, with nine pairs of chromosomes evident (Fig. 6C). Fewer than nine (five to eight) pairs of chromosomes have been observed in some instances, particularly in the D-26 strain of A. bisporus. The significance of such variation is not now known. Spindle pole bodies exist in higher fungi, and close association of nuclear chromatin with these structures has been reported (7, 8, 25, 33, 34). Depending on orientation of the spindle apparatus, spindle pole bodies, if indeed Feulgen positive, might be mistaken for chromosomes by fluorescent microscopy. This could be the basis for apparent variation in chromosome number in basidia of A.
VOL. 126, 1976
DEVELOPMENT OF AGARICUS BISPORUS
419
FIG. 1. Basidiospores of A. bisporus 70-2 stained with auramin 0 and examined by fluorescent micros(A) Spore sample showing variation in nuclear distribution. (B) Six-nucleated spore prior to germination. (C-E) Stages of spore germination showing migration of nuclei into the germ tube.
copy.
TABLE 1. Distribution of nuclei in spores of A. bisporusa Strain D-26
Strain 70-2
No. of nuclei
No. ofserved spores ob-
Percentage of sam-
No. of nuclei
No. of spores observed
Percentage of sample
0 1 2 3 4 5 6 7 8
17 0 75 229 234 36 19 8
2.8 0 12.2 37.1 37.9 5.8 3.1 1.3 0.7
0 1 2 3 4
25 2 82 69 276
5.4 0.4 17.6 14.7 59.3
5 6 7 8
6 4 1 1
1.3 0.9 0.2 0.2
a
4
ple
Based on counts of randomly selected spores in samples examined by fluorescent microscopy.
420
SAKSENA ET AL.
J. BACTERIOL.
FIG. 2. Scanning electron microscopy of basidiospores of A. bisporus. (A) Dormant spores en masse from an air-dried spore print. (B, C) Early stages of spore germination. (D) Germinated basidiospore; arrow indicates apiculus or point of spore attachment to basidium. (E, F) Later stages of spore germination showing an unbranched and a branched germ tube.
VOL. 126, 1976
DEVELOPMENT OF AGARICUS BISPORUS
421
FIG. 3. Mycelia of A. bisporus. (A) Fluorescent Feulgen-stained nuclei in a mycelial network; arrow indicates nucleus migrating between hyphae. (B) Mycelial network showing encrustation on some of hyphae. (C) Encrusted hypha bearing crystals of two morphological types.
,
gh
A
4 D
FIG. 4. Nuclei in hyphae of A. bisporus. (A) Elongated nuclei during mitosis; arrow inddicates nucleus with a "ttwo-track" configuration. (B) Branched hypha with nuclei in three configurations: co)ondensed (large arrow), cliffuse (small arrow), and elongated ("two-track"). (C, D) Trikaryotic cells. (E) Phase-contrast micrograjph of cell depicted in (D), showing relative advantage of the fluorescent Feulgen tec)hnique; arrows indicate Ipositions of three nuclei in the cell. 422
VOL. 126, 1976
DEVELOPMENT OF AGARICUS BISPORUS
423
FIG. 5. Mushrooms of A. bisporus D-26. (A) Normal mushrooms showing deep gills and prominent veil (arrow). (B, C) Abnormal or "open-veil" mushrooms, the extreme condition being shown in (C).
424
SAKSENA ET AL.
bisporus, but aneuploidy or actual variation in chromosome number cannot be ruled out until further studies are undertaken. Stages of meiosis subsequent to prophase I exhibit elongated configurations (Fig. 7A and B). Evidence for peripheral distribution of deoxyribonucleic acid in nuclei during meiotic interphase is shown in Fig. 7C. Typically, meiosis yields four products, which are distributed into two spores (Fig. 7D, 8A, and 9B, E, and F). Single-spored (Fig. 8B) and multispored (Fig. 9C and D) basidia are present in strains of A. bisporus as well. Strain D-26 and strain 70-2 do not appear to form four-spored basidia (Table 2). Gill surfaces of mushrooms in various stages of development were examined in detail by
J . B3ACTERIOL .
scanning electron microscopy (Fig. 9 and 10). Stigmata generally develop in parallel at the apex of the basidium. Basidia in different stages of maturation are often seen in one area of a gill tissue taken from an opened mushroom (Fig. 9A and 1OA). Young basidia, common in unopened mushrooms, are cylindrical (slightly swollen at the apex) and their surfaces are smooth. Subsequently, the basidial cells swell and their surfaces become slightly wrinkled. This wrinkled appearance persists throughout spore development. After spore discharge, basidia collapse and appear flaccid. Basidia with greater than three spores were not observed in either strain D-26 or 70-2 (Table 2). In strain 70-2, even three-spored basidia were extremely rare (Fig. 1OA and B). In sev-
'II-.-FIG. 6. Early stages of meiosis in A. bisporus 70-2. (A) Early prophase. (B) Middle prophase showing unpaired chromosomes. (C) Pachytene stage of meiosis I showing nine pairs of chromosomes.
FIG. 7. Meiosis in A. bisporus D-26. (A, B) Metaphase of meiosis I; arrow shows elongated or "two-track" configuration in the meiotic nucleus. (C) Late-telophase nuclei during meiosis. (D) Anaphase of meiosis II.
DEVELOPMENT OF AGARICUS BISPORUS
VOL. 126, 1976
1 pm
-
425
____
FIG. 8. Nuclei in basidia of A. bisporus D-26. (A) Meiotic tetrad in two-pronged basidium. (B) Singlespored basidium.
426
SAKSENA ET AL.
J. BACTERIOL.
-I..-s FIG. 9. Scanning electron microscopy of developing basidia in A. bisporus D-26. (A) Gill surface of mature mushroom. (B through D). Young two-spored and three-spored basidia. (E) Mature two-spored basidium. (F) Basidium after spore discharge.
DEVELOPMENT OF AGARICUS BISPORUS
VOL. 126, 1976
r-S7r
427
_r_
FIG. 10. Scanning electron microscopy offertile and sterile surfaces ofA. bisporus 70-2. (A, B) Portion of gills showing high frequencies of two-spored basidia. (C) Sterile structures on an aborted gill surface of an "open-veil" mushroom the same strain.
TABLE 2. Distribution of spores on basidia of A. bisporusa Strain 70-2
Strain D-26
No. of spores
No. Perof centage basi- of sam-
1 2
dia 15 724
No. of spores
No. of basi-
ple
2.0 93.6
1 2
Percent-
Perof age of
diaa 34 4.6 694 95.0
3 3 0.4 4.4 4 or more 0 0 0 a Based on counts from randomly selected areas of mushroom gills examined by scanning electron microscopy. 33 3 4 or more 0
eral instances, apparent four-spored basidia were seen at low magnification (x 1,000), but these, in fact, represented overlapping or distorted situations which could be resolved clearly through scanning electron microscopy either by tilting the specimen or by increasing the magnification. A high frequency of multispored basidia has been reported for some strains of A. bisporus (5, 26, 44). These studies, based on light microscopy, may not be quantitatively accurate in view of the limitations on resolution of structures on gill surfaces at low magnification. The absence of basidia with four or more spores, in our strains, suggests that considerable variation exists between various strains of A. bisporus. The surfaces of gill-less or "open-veil" mushrooms, also examined by scanning electron microscopy, appeared to be sterile (Fig. 10C). The nature or the cause of this condition has not yet been ascertained. ACKNOWLEDGMENTS This research was supported by the Butler County Mushroom Farm, Inc., Worthington, Pa., through a fellowship to the Carnegie-Mellon Institute of Research.
LITERATURE CITED 1. Brushaber, J. A., C. L. Wilson, and J. R. Aist. 1967. Asexual nuclear division of some plant pathogenic fungi. Phytopathology 57:43-46. 2. Cohen, A. L. 1973. Critical point drying, p. 44-112. In M. A. Hayat (ed.), Principles and techniques of scanning electron microscopy, vol. 1. Van Nostrand Reinhold Co., New York. 3. Colson, B. 1935. The cytology of the mushroom Psalliota campestris Quel. Ann. Bot. (London) 49:1-18. 4. Day, A. W. 1972. Genetic implications of current models of somatic nuclear division in fungi. Can. J. Bot. 50:1337-1347. 5. Elliot, T. J. 1972. Sex and the single spore. Mushroom Sci. 8:11-18. 6. Evans, H. J. 1956. Chromosomes of the cultivated mushroom. Nature (London) 178:1005-1006. 7. Girbardt, M. 1971. Ultrastructure of the fungal nucleus. II. The kinetochore equivalent (KCE). J. Cell Sci. 2:453-473. 8. Gull, K., and R. J. Newsam. 1975. Meiosis in basidiomycetous fungi. I. Fine structure of spindle pole body organization. Protoplasma 83:247-257. 9. Harris, J. L., H. B. Howe, and I. L. Roth. 1975. Scanning electron microscopy of surface and internal features of developing perithecia of Neurospora crassa. J. Bacteriol. 122:1239-1246. 10. Hashioka, Y. 1971. Scanning and transmission electron micrographs of shiitaki, Lentinus edodes (Berk.) Sing Rep. Tottori Mycol. Inst. 9:1-10. 11. Hashmi, M. H., B. Kendrick, and G. Morgan-Jones. 1972. Mitosis in three hyphomycetes. Can. J. Bot. 50:2375-2578. 12. Heath, I. B. 1974. Mitosis in the fungus Thraustotheca clavata. J. Cell Biol. 60:204-220. 13. Hou, H. H., and L. C. Wu. 1972. Nuclear behavior of cultivated mushroom. Bot. Bull. Acad. Sin. 13:82-91. 14. Huang, H. C., R. D. Tinline, and L. C. Fowke. 1975. Ultrastructure of somatic mitosis in a diploid strain of plant pathogenic fungus Cochliobolus sativus. Can. J. Bot. 53:403-414. 15. Hughes, D. T. 1961. Chromosomes of the wild mushroom. Nature (London) 190:285-286. 16. Jiti, H. 1965. Zytologische Studien uber die Gattung Agaricus. Mushroom Sci. 6:77-81. 17. Kligman, A. M. 1943. Some cultural and genetic problems in the cultivation of the mushroom Agaricus campestris. F. Am. J. Bot. 30:745-763. 18. Laane, M. M., and T. Lie. 1975. Examination of fungal nuclei with Feulgen fluorescence method. Mikroskopie 31:85-90. 19. Lambert, E. B. 1929. The production of normal sporo-
428
SAKSENA ET AL.
J. BACTERIOL.
phores in monosporous cultures of Agaricus bisporus. 310. Mycologia 21:333-335. 34. Raju, N. B., and B. C. Lu. 1973. Meiosis in Coprinus. 20. Lemke, P. A., J. R. Ellison, R. Marino, B. Morimoto, IV. Morphology and behavior of spindle pole bodies. E. Arons, and P. Kohman. 1975. Fluorescent Feulgen J. Cell Sci. 12:131-141. staining of fungal nuclei. Exp. Cell Res. 96:367-373. 35. Raper, C. A., J. R. Raper, and R. E. Miller. 1972. 21. Le Roux, P. 1968. Action du gaz carbonique sur le Genetic analysis of the life cycle ofAgaricus bisporus. metabolisme du carpophore dAgaricus bisporus. I. Mycologia 74:1088-1117. Les acides organiques et les acides amines libres. 36. Rasmussen, C. R., M. G. Amsen, and G. Holmgaard. Mushroom Sci. 7:31-36. 1962. "Open veiled" or "hard gilled" mushrooms. 22. Losel, D. M. 1964. The stimulation of spore germination Mushroom Sci. 4:416-429. in Agaricus bisporus by living mycelium. Ann. Bot. 37. Rast, D., and E. J. Stauble. 1970. On the mode of action 28:541-554. of isovaleric acid in stimulating the germination of 23. Lowry, R. J., and A. S. Sussman. 1968. Ultrastructural Agaricus bisporus spores. New Phytol. 69:557-566. changes during germination of ascospores of Neuro- 38. Robinow, C. F., and C. E. Caten. 1969. Mitosis in spora tetrasperma. J. Gen. Microbiol. 51:403-409. Aspergillus nidulans. J. Cell Sci. 5:403-431. 24. McKnight, K. H., and L. R. Batra. 1974. Scanning 39. San Antonio, J. P. 1971. A laboratory method to obtain electron microscopy in taxonomy of gyromitroid fruit from cased grain spawn of the cultivated mushfungi. Mich. Bot. 13:52-64. room, Agaricus bisporus. Mycologia 63:16-21. 25. McLaughlin, D. J. 1971. Centrosomes and microtubules 40. Sarazin, M. A. 1938. Evolution nucleaire de la baside during meiosis in the mushroom Boletus rubinellus. des basidiospores dans Agaricus campestris (var. culJ. Cell Biol. 50:737-745. tivee). C. R. Acad. Sci. Ser. D 206:275-278. 26. Miller, R. E. 1971. Evidence of sexuality in the culti- 41. Sarazin, M. A. 1939. Cultures monospermes dAgaricus vated mushroom, Agaricus bisporus. Mycologia campestris var. cultivee. C. R. Acad. Sci. Ser. D 63:630-634. 208:2015-2017. 27. Monocha, M. S. 1965. Fine structure of the Agaricus 42. Sass, J. E. 1928. A cytological study of a bispored form carpophore. Can. J. Bot. 43:1329-1333. of Psalliota campestris. Pap. Mich. Acad. Sci. Arts 28. Nakai, Y., and R. Ushiyama. 1974. Fine structure of Lett. 9:287-298. shiitaki, Lentinus edodes (Berk.) Sing. I. Scanning 43. Sass, J. E. 1936. Cytology of spore germination in the electron microscopy on basidia and basidiospores. bispored form of Psalliota campestris. Mycologia 28:431-432. Rep. Tottori Mycol. Inst. 11:1-6. 29. Newcomer, E. H. 1953. A new cytological and histologi- 44. Song, S. F., K. J. Hsu, and Y. L. Hsieh. 1972. Observations on the spored-basidium in the cultivated mushcal fixing fluid. Science 118:161. 30. Pegler, D. N., and T. W. K. Young. 1974. Critical point room (Agaricus bisporus). Mushroom Sci. 8:295-303. technique for scanning studies of basidiomycete hy- 45. Thielke, C. 1968. Die Substruktur der Zellen im menia. Trans. Br. Myco4. Soc. 63:175-211. Fruchtkorper von Psalliota bispora. Mushroom Sci. 31. Pelham, J. 1965. Techniques for mushroom genetics. 7:23-30. Mushroom Sci. 6:49-64. 46. Thielke, C. 1973. Intranuclear Mitosin in homokary32. Peng, J. T., and L. C. Wu. 1972. Variations in the otischen und dikaryotischen Mycelien der Basidiomycultivated mushroom, Agaricus bisporus. Mushroom ceten. Arch. Mikrobiol. 49:341-350. Sci. 8:103-113. 47. Vogel, F. S., and R. F. Weaver. 1972. Concerning the 33. Radu, M., R. Steinlauf, and Y. Koltin. 1974. Meiosis in induction of dormancy in spores of Agaricus bisporus. Schizophyllum commune. Arch. Microbiol. 98:301Exp. Cell Res. 75:95-104.
JOURNAL OF BACTERIOLOGY, Apr. 1976, p. 417-428 Copyright ©D 1976 American Society for Microbiology
Study on Development of Agaricus bisporus by Fluorescent Microscopy and Scanning Electron Microscopy K. N. SAKSENA,* R. MARINO, M. N. HALLER, AND P. A. LEMKE Carnegie-Mellon Institute of Research, Carnegie-Mellon University, Pittsburgh, Pennsylvania 15213
Received for publication 30 December 1975
Two strains of Agaricus bisporus have been investigated by fluorescent microscopy and scanning electron microscopy. Somatic nuclei, stained with auramin 0 and examined by fluorescent microscopy, appear to be randomly distributed, divide asynchronously, and assume a nonclassical or "two-track" configuration during mitotic metaphase. A similar configuration has been observed for nuclei during meiosis, but early meiosis in A. bisporus appears to be classical, usually with nine pairs of chromosomes evident during prophase I. Scanning electron microscopy has been used to document developmental stages in the formation and germination of basidiospores. Two-spored basidia were predominant, but occasionally one- or three-spored forms were observed. Fourspored basidia were absent, and uninucleated basidiospores were exceedingly rare to absent.
Studies on the cytology and genetics ofAgaricus bisporus (Lange) Imbach have shown consistently that the sexual cycle of this fungus is highly disorganized and atypical of a basidiomycete. The fungus has no semblance of dikaryotic organization, and the basidiospores, produced mostly two per basidium, contain an indefinite number of nuclei and represent predominantly self-fertile progeny (3, 5, 6, 13, 1517, 19, 20, 26, 27, 31, 32, 35, 40-45). We have examined the cytology and development of two strains of A. bisporus. Critical examination of these features has been possible through recent advances in fluorescent microscopy of fungal nuclei (18, 20) and through improvements in the preparation of biological specimens for scanning electron microscopy (2, 9, 10, 23, 24, 28, 30). MATERIALS AND METHODS Strains and media. Strains of A. bisporus designated D-26 and 70-2 were investigated. These strains represent, respectively, golden-white and light-brown varieties of the cultivated mushroom. Cultures were grown on a defined medium containing: asparagine, 1 g; glucose, 20 g; MgSO4 7 H20, 0.5 g; K2HPO4, 1 g; KH2PO4, 0.46 g; CaSO4, 0.5 g; FeCl3, 10 mg; thiamine, 120 Ag; distilled water, 1,000 ml; and agar, 20 g. Mushrooms were obtained commercially (Butler County Mushroom Farm, Inc., Worthington, Pa.) or were grown in the laboratory, using cased rye grain as a substrate (39). Basidiospores were collected as spore prints at 20 C and germinated either in the presence of isoamyl alcohol or isovaleraldehyde (1 and 0.1%, respectively, added to defined medium after autoclaving) or in the prox-
imity of a growing mycelium (23, 37, 47). Spores and mycelia were generally inoculated onto 5 cm2 membranes (DuPont PD 150 film) spread upon the surface of the defined medium. Cultures were incubated at 25 C, and membranes with adherent cultures were removed from the medium and prepared for microscope examination. Fluorescent microscopy. Procedures for examination of fungal nuclei by fluorescent Feulgen staining were described previously (20). Two dyes, auramin 0 (no. 10-120, Chroma-Gesellshaft, Stuttgart) and acrinol (no. Dl 660-6, Aldrich, Milwaukee), were used. Acrinol was used only to stain nuclei of basidial cells. Gill tissue was treated with Newcomer fixative (29) for at least 6 h and stored in 70% isopropanol prior to staining with the acrinol. In all other instances, fixation of cells was avoided and auramin 0 was used as the stain. Cells were placed in 1 N HCl at 60 C for at least 2 min prior to staining. Staining solutions were freshly prepared and contained 20 ml of water, 0.1 g of auramin 0 (or 0.1 g of acrinol), 0.2 g of potassium metabisulfite (0.2 g of sodium metabisulfite substituted if acrinol used), and 2 ml of 1 N HCl. Cells were stained for 20 min and then washed repeatedly in water and finally in 45% acetic acid. In each step of the procedure, cells were recovered from suspension by centrifugation (1,000 x g). Cells were placed on a microscope slide and air dried. The slide was then dipped momentarily in 95% ethanol, and material remaining on the slide was mounted for microscope examination in 65% sucrose (immersion oil substituted if acrinol used). A Leitz Orthoplan microscope fitted for incidentlight fluorescence with a Ploem vertical illuminator was used. The light source was an Osram superpressure mercury lamp (HBO 100 W/1 DC). During examination of material stained with auramin 0, the 117
418
SAKSENA ET AL.
microscope was set at position 3 with the following: one red suppression filter (BG38), two excitation filters (KP490), and one dichroic mirror (K490); no barrier filter was used. A X100 fluorite objective (1.39 numerical aperture) fitted with an iris (C. Reichert) was used. During examination of material stained with acrinol, the microscope was set at position 1 with the following: one excitation filter (UG1), one dichroic mirror (K400), and a barrier filter (K420). Either a X95 fluorite objective (1.32 numerical aperture) or a X90 apochromatic objective (1.40 numerical aperture) was used. Photographs were taken on Kodak Panatomic X film. Scanning electron microscopy. Segments of gill tissue (3 mm2) cut from fresh mushrooms at different stages of development were fixed with 6.25% glutaraldehyde in 0.05 M phosphate buffer (pH 7.0) for 3 h, washed repeatedly in buffer, and postfixed in 1% osmium tetraoxide for 1 h. After the washing, dehydration in a graded series of ethanol (30, 50, 70, and 90% for 15 min each, and 100% with three changes each of 15-min duration), and subsequent substitution in isoamylacetate for 15 min, critical point drying with liquid CO2 was performed with a Sorvall critical point dryer. Spore prints and cultures on membranes were air dried without fixation. Dried samples were coated with gold-palladium and examined with a JOELCO-JSM2 scanning electron microscope operating at 25 kV. The microscope was equipped with a nondispersive X-ray analyzer for elemental chemical analysis. Photographs were taken on Polaroid P/N55 film.
RESULTS AND DISCUSSION Initial experiments on the cytology of A. bisporus were conducted with basidiospores. These spores were collected as spore prints from opened mushrooms of two strains, D-26 and 702. Nuclei in these cells were stained with auramin 0 (Fig. 1A). The spores from the two strains proved to vary somewhat in nuclear distribution (Table 1). Uninucleated spores were observed rarely and only in one strain, 702. The majority of spores from this strain contained four nuclei, whereas the majority of spores derived from strain D-26 contained three or four nuclei. The surface of basidiospores examined by scanning electron microscopy proved to be smooth even at high (x 10,000) magnification (Fig. 2A). These dormant spores (47) can be stimulated to germinate at up to 50% frequency if placed in proximity to a growing mycelium of A. bisporus (22). Alternatively, spores can be induced to germinate by the addition of 1% isoamyl alcohol or 0.1 % isovaleraldehyde to the growth medium (37). Regardless of the method used to induce germination, spores do not germinate synchronously, and they produce a single germ tube per spore (Fig. 2B through F).
J. BACTERIOL.
The entire contents of a spore empty into this germ tube (Fig. 1B through E). Nuclei in mycelia do not divide synchronously or establish a dikaryon. Somatic nuclei are irregularly distributed and can be transferred to adjacent cells by hyphal anastomosis (Fig. 3A). Mature hyphae are often heavily encrusted (Fig. 3B and C). This encrustation, when observed by scanning electron microscopy, was resolved as crystals of at least three forms. X-ray analysis of crystals indicated the presence of potassium and calcium ions therein, and organic acids such as oxalate are assumed to be the major anionic component (21). Somatic nuclei of A. bisporus are often elongated (Fig. 4A) and assume a nonclassical configuration during mitotic division. This configuration has been observed in other higher fungi (1, 11, 12, 14, 38, 46) and has been called a "twotrack" configuration (4). Asynchrony of nuclei in hyphae of A. bisporus is evident in Fig. 4B, which shows a single, branched cell containing three types of nuclei: condensed, diffuse, and elongated ("two-track"). In A. bisporus the most common number of nuclei per vegetative cell is 3 (Fig. 4C through E), but as many as 25 nuclei per cell have been counted. Greater than 25 nuclei per cell have been seen in inflated (nonvegetative) cells found in the cap and stipe of mushrooms. Young mushrooms with normal morphology (Fig. 5A) have deep gills and a prominent veil. At maturity these mushrooms produce from one to several billion basidiospores. Abnormal mushrooms with reduced gill formation and scant veil tissue are sometimes found (Fig. 5B). In an extreme condition, asporogenic or imperfect mushrooms are formed (Fig. 5C), a condition that has been termed "open veil" (36). Basidia of A. bisporus develop asynchronously, and details of their meiosis are shown in Fig. 6 and 7. Nuclei and chromosomes in these cells were stained with acrinol. Early meiosis or prophase I appears to be classical, with nine pairs of chromosomes evident (Fig. 6C). Fewer than nine (five to eight) pairs of chromosomes have been observed in some instances, particularly in the D-26 strain of A. bisporus. The significance of such variation is not now known. Spindle pole bodies exist in higher fungi, and close association of nuclear chromatin with these structures has been reported (7, 8, 25, 33, 34). Depending on orientation of the spindle apparatus, spindle pole bodies, if indeed Feulgen positive, might be mistaken for chromosomes by fluorescent microscopy. This could be the basis for apparent variation in chromosome number in basidia of A.
VOL. 126, 1976
DEVELOPMENT OF AGARICUS BISPORUS
419
FIG. 1. Basidiospores of A. bisporus 70-2 stained with auramin 0 and examined by fluorescent micros(A) Spore sample showing variation in nuclear distribution. (B) Six-nucleated spore prior to germination. (C-E) Stages of spore germination showing migration of nuclei into the germ tube.
copy.
TABLE 1. Distribution of nuclei in spores of A. bisporusa Strain D-26
Strain 70-2
No. of nuclei
No. ofserved spores ob-
Percentage of sam-
No. of nuclei
No. of spores observed
Percentage of sample
0 1 2 3 4 5 6 7 8
17 0 75 229 234 36 19 8
2.8 0 12.2 37.1 37.9 5.8 3.1 1.3 0.7
0 1 2 3 4
25 2 82 69 276
5.4 0.4 17.6 14.7 59.3
5 6 7 8
6 4 1 1
1.3 0.9 0.2 0.2
a
4
ple
Based on counts of randomly selected spores in samples examined by fluorescent microscopy.
420
SAKSENA ET AL.
J. BACTERIOL.
FIG. 2. Scanning electron microscopy of basidiospores of A. bisporus. (A) Dormant spores en masse from an air-dried spore print. (B, C) Early stages of spore germination. (D) Germinated basidiospore; arrow indicates apiculus or point of spore attachment to basidium. (E, F) Later stages of spore germination showing an unbranched and a branched germ tube.
VOL. 126, 1976
DEVELOPMENT OF AGARICUS BISPORUS
421
FIG. 3. Mycelia of A. bisporus. (A) Fluorescent Feulgen-stained nuclei in a mycelial network; arrow indicates nucleus migrating between hyphae. (B) Mycelial network showing encrustation on some of hyphae. (C) Encrusted hypha bearing crystals of two morphological types.
,
gh
A
4 D
FIG. 4. Nuclei in hyphae of A. bisporus. (A) Elongated nuclei during mitosis; arrow inddicates nucleus with a "ttwo-track" configuration. (B) Branched hypha with nuclei in three configurations: co)ondensed (large arrow), cliffuse (small arrow), and elongated ("two-track"). (C, D) Trikaryotic cells. (E) Phase-contrast micrograjph of cell depicted in (D), showing relative advantage of the fluorescent Feulgen tec)hnique; arrows indicate Ipositions of three nuclei in the cell. 422
VOL. 126, 1976
DEVELOPMENT OF AGARICUS BISPORUS
423
FIG. 5. Mushrooms of A. bisporus D-26. (A) Normal mushrooms showing deep gills and prominent veil (arrow). (B, C) Abnormal or "open-veil" mushrooms, the extreme condition being shown in (C).
424
SAKSENA ET AL.
bisporus, but aneuploidy or actual variation in chromosome number cannot be ruled out until further studies are undertaken. Stages of meiosis subsequent to prophase I exhibit elongated configurations (Fig. 7A and B). Evidence for peripheral distribution of deoxyribonucleic acid in nuclei during meiotic interphase is shown in Fig. 7C. Typically, meiosis yields four products, which are distributed into two spores (Fig. 7D, 8A, and 9B, E, and F). Single-spored (Fig. 8B) and multispored (Fig. 9C and D) basidia are present in strains of A. bisporus as well. Strain D-26 and strain 70-2 do not appear to form four-spored basidia (Table 2). Gill surfaces of mushrooms in various stages of development were examined in detail by
J . B3ACTERIOL .
scanning electron microscopy (Fig. 9 and 10). Stigmata generally develop in parallel at the apex of the basidium. Basidia in different stages of maturation are often seen in one area of a gill tissue taken from an opened mushroom (Fig. 9A and 1OA). Young basidia, common in unopened mushrooms, are cylindrical (slightly swollen at the apex) and their surfaces are smooth. Subsequently, the basidial cells swell and their surfaces become slightly wrinkled. This wrinkled appearance persists throughout spore development. After spore discharge, basidia collapse and appear flaccid. Basidia with greater than three spores were not observed in either strain D-26 or 70-2 (Table 2). In strain 70-2, even three-spored basidia were extremely rare (Fig. 1OA and B). In sev-
'II-.-FIG. 6. Early stages of meiosis in A. bisporus 70-2. (A) Early prophase. (B) Middle prophase showing unpaired chromosomes. (C) Pachytene stage of meiosis I showing nine pairs of chromosomes.
FIG. 7. Meiosis in A. bisporus D-26. (A, B) Metaphase of meiosis I; arrow shows elongated or "two-track" configuration in the meiotic nucleus. (C) Late-telophase nuclei during meiosis. (D) Anaphase of meiosis II.
DEVELOPMENT OF AGARICUS BISPORUS
VOL. 126, 1976
1 pm
-
425
____
FIG. 8. Nuclei in basidia of A. bisporus D-26. (A) Meiotic tetrad in two-pronged basidium. (B) Singlespored basidium.
426
SAKSENA ET AL.
J. BACTERIOL.
-I..-s FIG. 9. Scanning electron microscopy of developing basidia in A. bisporus D-26. (A) Gill surface of mature mushroom. (B through D). Young two-spored and three-spored basidia. (E) Mature two-spored basidium. (F) Basidium after spore discharge.
DEVELOPMENT OF AGARICUS BISPORUS
VOL. 126, 1976
r-S7r
427
_r_
FIG. 10. Scanning electron microscopy offertile and sterile surfaces ofA. bisporus 70-2. (A, B) Portion of gills showing high frequencies of two-spored basidia. (C) Sterile structures on an aborted gill surface of an "open-veil" mushroom the same strain.
TABLE 2. Distribution of spores on basidia of A. bisporusa Strain 70-2
Strain D-26
No. of spores
No. Perof centage basi- of sam-
1 2
dia 15 724
No. of spores
No. of basi-
ple
2.0 93.6
1 2
Percent-
Perof age of
diaa 34 4.6 694 95.0
3 3 0.4 4.4 4 or more 0 0 0 a Based on counts from randomly selected areas of mushroom gills examined by scanning electron microscopy. 33 3 4 or more 0
eral instances, apparent four-spored basidia were seen at low magnification (x 1,000), but these, in fact, represented overlapping or distorted situations which could be resolved clearly through scanning electron microscopy either by tilting the specimen or by increasing the magnification. A high frequency of multispored basidia has been reported for some strains of A. bisporus (5, 26, 44). These studies, based on light microscopy, may not be quantitatively accurate in view of the limitations on resolution of structures on gill surfaces at low magnification. The absence of basidia with four or more spores, in our strains, suggests that considerable variation exists between various strains of A. bisporus. The surfaces of gill-less or "open-veil" mushrooms, also examined by scanning electron microscopy, appeared to be sterile (Fig. 10C). The nature or the cause of this condition has not yet been ascertained. ACKNOWLEDGMENTS This research was supported by the Butler County Mushroom Farm, Inc., Worthington, Pa., through a fellowship to the Carnegie-Mellon Institute of Research.
LITERATURE CITED 1. Brushaber, J. A., C. L. Wilson, and J. R. Aist. 1967. Asexual nuclear division of some plant pathogenic fungi. Phytopathology 57:43-46. 2. Cohen, A. L. 1973. Critical point drying, p. 44-112. In M. A. Hayat (ed.), Principles and techniques of scanning electron microscopy, vol. 1. Van Nostrand Reinhold Co., New York. 3. Colson, B. 1935. The cytology of the mushroom Psalliota campestris Quel. Ann. Bot. (London) 49:1-18. 4. Day, A. W. 1972. Genetic implications of current models of somatic nuclear division in fungi. Can. J. Bot. 50:1337-1347. 5. Elliot, T. J. 1972. Sex and the single spore. Mushroom Sci. 8:11-18. 6. Evans, H. J. 1956. Chromosomes of the cultivated mushroom. Nature (London) 178:1005-1006. 7. Girbardt, M. 1971. Ultrastructure of the fungal nucleus. II. The kinetochore equivalent (KCE). J. Cell Sci. 2:453-473. 8. Gull, K., and R. J. Newsam. 1975. Meiosis in basidiomycetous fungi. I. Fine structure of spindle pole body organization. Protoplasma 83:247-257. 9. Harris, J. L., H. B. Howe, and I. L. Roth. 1975. Scanning electron microscopy of surface and internal features of developing perithecia of Neurospora crassa. J. Bacteriol. 122:1239-1246. 10. Hashioka, Y. 1971. Scanning and transmission electron micrographs of shiitaki, Lentinus edodes (Berk.) Sing Rep. Tottori Mycol. Inst. 9:1-10. 11. Hashmi, M. H., B. Kendrick, and G. Morgan-Jones. 1972. Mitosis in three hyphomycetes. Can. J. Bot. 50:2375-2578. 12. Heath, I. B. 1974. Mitosis in the fungus Thraustotheca clavata. J. Cell Biol. 60:204-220. 13. Hou, H. H., and L. C. Wu. 1972. Nuclear behavior of cultivated mushroom. Bot. Bull. Acad. Sin. 13:82-91. 14. Huang, H. C., R. D. Tinline, and L. C. Fowke. 1975. Ultrastructure of somatic mitosis in a diploid strain of plant pathogenic fungus Cochliobolus sativus. Can. J. Bot. 53:403-414. 15. Hughes, D. T. 1961. Chromosomes of the wild mushroom. Nature (London) 190:285-286. 16. Jiti, H. 1965. Zytologische Studien uber die Gattung Agaricus. Mushroom Sci. 6:77-81. 17. Kligman, A. M. 1943. Some cultural and genetic problems in the cultivation of the mushroom Agaricus campestris. F. Am. J. Bot. 30:745-763. 18. Laane, M. M., and T. Lie. 1975. Examination of fungal nuclei with Feulgen fluorescence method. Mikroskopie 31:85-90. 19. Lambert, E. B. 1929. The production of normal sporo-
428
SAKSENA ET AL.
J. BACTERIOL.
phores in monosporous cultures of Agaricus bisporus. 310. Mycologia 21:333-335. 34. Raju, N. B., and B. C. Lu. 1973. Meiosis in Coprinus. 20. Lemke, P. A., J. R. Ellison, R. Marino, B. Morimoto, IV. Morphology and behavior of spindle pole bodies. E. Arons, and P. Kohman. 1975. Fluorescent Feulgen J. Cell Sci. 12:131-141. staining of fungal nuclei. Exp. Cell Res. 96:367-373. 35. Raper, C. A., J. R. Raper, and R. E. Miller. 1972. 21. Le Roux, P. 1968. Action du gaz carbonique sur le Genetic analysis of the life cycle ofAgaricus bisporus. metabolisme du carpophore dAgaricus bisporus. I. Mycologia 74:1088-1117. Les acides organiques et les acides amines libres. 36. Rasmussen, C. R., M. G. Amsen, and G. Holmgaard. Mushroom Sci. 7:31-36. 1962. "Open veiled" or "hard gilled" mushrooms. 22. Losel, D. M. 1964. The stimulation of spore germination Mushroom Sci. 4:416-429. in Agaricus bisporus by living mycelium. Ann. Bot. 37. Rast, D., and E. J. Stauble. 1970. On the mode of action 28:541-554. of isovaleric acid in stimulating the germination of 23. Lowry, R. J., and A. S. Sussman. 1968. Ultrastructural Agaricus bisporus spores. New Phytol. 69:557-566. changes during germination of ascospores of Neuro- 38. Robinow, C. F., and C. E. Caten. 1969. Mitosis in spora tetrasperma. J. Gen. Microbiol. 51:403-409. Aspergillus nidulans. J. Cell Sci. 5:403-431. 24. McKnight, K. H., and L. R. Batra. 1974. Scanning 39. San Antonio, J. P. 1971. A laboratory method to obtain electron microscopy in taxonomy of gyromitroid fruit from cased grain spawn of the cultivated mushfungi. Mich. Bot. 13:52-64. room, Agaricus bisporus. Mycologia 63:16-21. 25. McLaughlin, D. J. 1971. Centrosomes and microtubules 40. Sarazin, M. A. 1938. Evolution nucleaire de la baside during meiosis in the mushroom Boletus rubinellus. des basidiospores dans Agaricus campestris (var. culJ. Cell Biol. 50:737-745. tivee). C. R. Acad. Sci. Ser. D 206:275-278. 26. Miller, R. E. 1971. Evidence of sexuality in the culti- 41. Sarazin, M. A. 1939. Cultures monospermes dAgaricus vated mushroom, Agaricus bisporus. Mycologia campestris var. cultivee. C. R. Acad. Sci. Ser. D 63:630-634. 208:2015-2017. 27. Monocha, M. S. 1965. Fine structure of the Agaricus 42. Sass, J. E. 1928. A cytological study of a bispored form carpophore. Can. J. Bot. 43:1329-1333. of Psalliota campestris. Pap. Mich. Acad. Sci. Arts 28. Nakai, Y., and R. Ushiyama. 1974. Fine structure of Lett. 9:287-298. shiitaki, Lentinus edodes (Berk.) Sing. I. Scanning 43. Sass, J. E. 1936. Cytology of spore germination in the electron microscopy on basidia and basidiospores. bispored form of Psalliota campestris. Mycologia 28:431-432. Rep. Tottori Mycol. Inst. 11:1-6. 29. Newcomer, E. H. 1953. A new cytological and histologi- 44. Song, S. F., K. J. Hsu, and Y. L. Hsieh. 1972. Observations on the spored-basidium in the cultivated mushcal fixing fluid. Science 118:161. 30. Pegler, D. N., and T. W. K. Young. 1974. Critical point room (Agaricus bisporus). Mushroom Sci. 8:295-303. technique for scanning studies of basidiomycete hy- 45. Thielke, C. 1968. Die Substruktur der Zellen im menia. Trans. Br. Myco4. Soc. 63:175-211. Fruchtkorper von Psalliota bispora. Mushroom Sci. 31. Pelham, J. 1965. Techniques for mushroom genetics. 7:23-30. Mushroom Sci. 6:49-64. 46. Thielke, C. 1973. Intranuclear Mitosin in homokary32. Peng, J. T., and L. C. Wu. 1972. Variations in the otischen und dikaryotischen Mycelien der Basidiomycultivated mushroom, Agaricus bisporus. Mushroom ceten. Arch. Mikrobiol. 49:341-350. Sci. 8:103-113. 47. Vogel, F. S., and R. F. Weaver. 1972. Concerning the 33. Radu, M., R. Steinlauf, and Y. Koltin. 1974. Meiosis in induction of dormancy in spores of Agaricus bisporus. Schizophyllum commune. Arch. Microbiol. 98:301Exp. Cell Res. 75:95-104.
